Microalgae Culture Protocol

t Microalgae Culture Protocol

Aquaculture Division Harbor Branch OceanographicInstitutionInc. t

MICROALGAE CULTURE PROTOCOL Aquaculture Division HARBOR BRANCH OCEANOGRAPHIC INSTITUTION INC.

Juan C. Jaramillo Harbor Branch Oceanographic Inst. 5600 U.S. I North Flo Pierce, Florida 34946

Key words: Sterile techniques, microalgae, nutrients, growth.

Abstract

The protocol for culturing microalgae at the Aquaculture Division of Harbor Branch Oceanographic Institution is explained in detail. The purpose of this document is to serve as a step-by-step guide for technicians in the microalgae laboratory. Sterile techniques, culture vessel cleaning and preparation, nutrient and media preparation, inoculations, and culture room maintenance are described. Systems diagrams and flow charts indicating culture techniques and paths are included. Appendices on autoclaving, filtration and chemical reagents preparation are also included.

Table of Contents Page Introduction 1 1.0 Stock Cultures (10 ml) 2 1.1 Clones Used at HBOI Aquaculture 2 1.1.1 Tiso (Isochrysis galbana) 3 1.1.2 Cg (Chaetoceros gracilis) 3 1.1.3 Tw iTh.alassiosira weissflogiiy 3 1.2 Culture Vessel Preparation 3 1.3 Media Preparation 4 1.4 Transfers 6 1.5 Stock Culture Table 7 2.0 Small-scale Cultures (500-6000 ml) 7 2.1 Culture Vessel Preparation 8 2.2 Culture Media Preparation 8 2.3 Inoculation 9 2.4 Small-scale Culture Table 10 3.0 Carboy Cultures (16-20 liters) 10 3.1 Culture Vessel Preparation 1 1 3.2 Culture Media Preparation 12 3.3 Inoculations 14 3.4 Culture Room 14 4.0 Cylinder Cultures in Room #2 (185- 208 liters) 16 4.1 Culture Vessel Preparation 17 4.2 Culture Media Preparation 1 8 4.3 Inoculation 19 4.4 Harvest 20 4.5 Culture Room 20 6.0 Algae Greenhouse Cultures 21 5.1 Culture Vessel Preparation 2 1 5.2 Culture Media Preparation 22 5.3 Inoculations 23 5.4 Greenhouses 23 Appendix Autoclaving 25 Appendix II Filtration and U.V. System 30 Appendix III Thiosulfate and Silicates 32

At the Aquaculture Division only few species of microalgae are used to feed our organisms. The most important is the small flagellate Isochrysis g albana which is used to feed larval as well as juvenile and adult clams. The second most important species in our lab is Chaetoceros gracilis, a small diatom used mostly to feed juvenile and adult stages of clams and larval stages of shrimp and other invertebrates. Also used for the same purpose is the diatom Th alassiosira w eissflogii. In addition to these species, the lab cultures Ellipsodon sp., Tet raselmis sp., Nannocloropsis sp., and Chlorella sp., to feed clams, rotifers, artemia and ornamental larval shrimp. Other clones not mentioned here are maintained in our small collection but they are rarely used for feedings.

Of utmost importance to the operation of growing microalgae in our laboratory is the proper utilization of Sterile Techniques in every step of the process in the first stages of batch cullure (inside laboratory, rooms #1 and #2). Without care taken in handling containers, media and instruments, you are likely to get contamination soon after you order new stock cultures. Those cultures are not axenic I to begin with (axenic cultures are a myth, really l), but they are pretty clean, and will last for years without a problem if precautions are taken. Some people prefer to move cultures fast through the laboratory to avoid "crashes">, I prefer to move slower, let things get dense enough, and be very meticulous with the technique to avoid those crashes. That way you are never in a rush and you do not have to live stressed and on "the edge" all the time.

Our operation starts with 10 ml sock cultures that are progressively scaled up to about 3000 liters. This guide takes you from the very beginning of the process to the stage where algae is ready to be fed to the animals in our facility.

1 Cultures with only one species of any microorganism. If a culture has bacteria, but still survives well, it is called "monoalgal culture".

2 Term used when a culture abruptly dies due to contamination or nutrient depletion. Generally a crash involves decoloration of the culture, the presence of clumps and sometimes a foul smell.

Stock cultures are those used to start new batches of cultured algae for the laboratory. They are obtained by two different means. One is the use of a technique, called Single Cell Isolation from water samples collected in the field. This technique is time consuming, difficult to master and the results are unreliable, therefore, it is not been used at Harbor Branch. The best way to obtain good quality stock cultures is to buy them from other laboratories. We use two main sources: The University of Texas (UTEX), and Bigelow Laboratories (CCMP). The collection at UTEX is extensive and the clones obtained from them in the last years have been very reliable. They are also inexpensive ($10 per test tube). CCMP also has a good collection but the cultures are considerably more expensive ($40-$100 per test tube). Cultures can also be obtained from the American Type Culture Collection (see addresses below).

1.1 Clones Used at HBOI Aquaculture

The clones used at HBOI can be ordered by letter, phone, fax, or usmg the internet at:

Culture Collection of Algae (UTEX) Department of Botany The University of Texas at Austin Austin, Tx 78713-7460 Tel: (512) 471-4019 Fax: (512) 471-3878 URL: http://www .botany.utexas.edu/i nfores/utex/

National Center for Culture of Marine Phytoplankton (CCMP) Bigelow Laboratory for Ocean Sciences West Boothbay Harbor, Maine 04575 Tel: (207) 633-9600 Fax: (207) 633-9641

American Type Culture Collection 1230I Parkland Drive Rockville, Maryland 20852-1776 Tel: (800) 638-6597 Fax: (301) 231-5826

We have one clone of Tiso from the UTEX collection. Its code is: LB2307

Size: 4-7 urn x 4 urn Doublings day-t: 2.89 TO Range: 16-34°C (28°C optimum)

1.1.2 Cg (Chaetoceros gracilis)

We have one clone of Cg from the UTEX collection. Its code is: LB 2375

Size: 5-7 urn x 4 urn Doublings day-i: 4.3 TO Range: Not determined (28-32°C optimum)

1.1.3 Tw tTh.alassiosira weissflogii)

We have one clone of Tw also, called 'Actin,' from The University of Miami. This clone can also be obtained from CCMP.

Size: 12-24 um diameter Doublings day-r: 1.39 TO Range: IO-30°C (25-30°C optimum)

When cultures are ordered make the people at Receiving aware of the shipment so they do not put them in the freezer or leave them exposed to the sun. Once in the lab, place them in a rack on the culture table and wait a day or two before you transfer them to new test tubes.

1.2 Culture Vessel Preparation

Stock cultures come from the providers in glass or plastic test tubes (10 ml). They are transferred into borosilicate test tubes with lose caps or lids. Tubes must be washed even if they are just out of the box. To wash tubes follow the next steps:

• Rinse them very well with Reverse Osmosis (R.O.) water several times. The last rinse is done using distilled water or deionized water. It is very important that all traces of soap are removed from inside the tubes.

• Pour 10 or 15 ml of D.l. or distilled water in each tube. This is done to trap copper residues left by the autoclaving process (The autoclave has copper plumbing) that can eventually kill the microalgae.

• Autoclave tubes for 30 minutes at 125°C (see the "Autoclaving" appendix).

• Let tubes cool down for 24 hr and store them In a dark, cool clean place until use.

1.3 Media Preparation

We buy nutrient stocks ("algae food" or f/2) rather than prepare them from chemical reagents in our lab, mainly to save time. They are bought in 5 gallon containers and re-bottled in smaller teflon or plastic containers for (internal) indoor use. We order nutrients from:

Aquacenter Tel: (800) 748-8921 Fax: (601) 378-2862/378-2861

There are two solutions to buy:

Solution A (trace metals / catalog # 1081 OA) Solution B (P, N and vitamins / catalog # 1081OB)

To sterilize media for the stock cultures, seawater must be passed through a series of filters ranging from 25 urn to 3 um, then through U.V., and finally through a fine 0.2 urn filter (Nalgenew, Fig 1.)

To do this make sure the filtration system is working and the U.V. lights are turned on (see the "Filtration and U.V. System" appendix). Filter water into a clean 500 ml Pyrex® glass container with a screw cap (Fig 2). Autoclave with the cap loose for 30 to 40 min. Let water cool down for 24 hr. Tighten the cap only when container is at room temperature, and store in dark, cool place until use.

Figure 2. 500 ml Pyrex'v glass container with a screw cap

Nutrients are re-bottled in small (500-1000 ml) dark teflon or polypropylene containers. Do not use polyethylene bottles, they will melt in the autoclave! Apply autoclaving indicator tape (the kind with white stripes across that become black when autoclaved) to the outside of each bottle and autoclave for 30 to 40 minutes. Wash hands with alcohol before handling tubes and pipettes. Using disposable sterile Pasteurw pipettes with cotton plugs, put 1 or 2 drops of sterile nutrient solution A and solution B in 500 ml of sterile filtered seawater. Use only one pipette per solution and container. Discard the D.1. water that was left in each tube when autoclaved. Under the laminar flow hood (Fig. 3), and with the help of a re-pipcttor or the Dispensette" syringe (previously sterilized) distribute 10 ml of media into an empty test tube, re-cap and place it in a 36-test tube rack. Repeat the procedure until you have empty the 500 ml container.

500 ml can roughly fill 1.5 racks (50 tubes). Make sure you add one or two drops of silicate solution to 500 ml of sterile seawater before you distribute media to the test tubes to be used for diatoms (see the "Thiosulfate and Silicates" appendix). Usually, one rack of tubes for diatoms is prepared for each two racks of regular, non-diatom, culture tubes. Store III a dark, cool, clean place until use.

1.4 Transfers

All transfers are done under the laminar flow hood (remember to turn it on before use l). Transfers have to be done every two to three weeks so algae does not die in the tubes from nutrient depletion. To transfer, clean your hands with alcohol, get an empty test tube rack, a rack with test tubes with sterile media, and the culture rack (Fig. 4) .

. .

Figure 4. Culture test tube racks

Place them under the hood and using Pasteurw pipettes (previously sterilized) get 1 ml from a culture tube that has been placed momentarily in the vortex to mix it, and drop it in the new test tube. Discard the pipette (recycled pipettes are placed in a 1000 ml beaker with 10% Hel, but they can also be disposed of properly in the "glass trash" container). Repeat this operation for each tube always using a new pipette when transferring. If the pipette is dropped, or if it touches anything outside the test tubes,

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 6 discard both the pipette and the tubes to avoid any possibility of bacterial contamination. If possible, use a mask for your nose and mouth when transferring. Write the date on the record card of the cultures and in the log book, and place the newly transferred cultures and the back-up cultures on the culture table (Fig. 5).

srocx CUlTURES f-<"

Figure 5. Stock culture table showing shade screen between lamps and cultures

1.5 Stock Culture Table

The culture table must always have two layers of screen between the lights and the cultures and the thermometer hanging in the middle should show 22-26°C. If necessary, use a small fan to cool down the pocket of air between the lights and the plexiglass. Make sure to replace any bulb that does not work or that is blinking.

2.0 Small-scale Cultures (500-6000 ml)

What we call Small-scale Cultures are those of no more than 8 liters that can be kept on the culture table. Typically we keep such cultures in clear polycarbonate containers that can be autoclaved. We have 3 sizes: 1000 ml, 4 liters and 8 liters (Fig. 6).

Figure 6. Clear polycarbonate culture huckets

Polycarbonate buckets have to be washed with Alconox® detergent and a sponge to remove any organic matter and bacteria from previous cultures. Even if buckets are new, they have to be thoroughly cleaned before using them to culture microalgae. After the Alconox® wash, rinse the vessel with R.O. water to remove soap and leave some water in the bucket to trap copper during the autoclaving process. Clean the lids with Alconox® soap also, and rinse them well with R.O. water. Assemble the containers put some autoclave indicator tape. Autoclave for 40 minutes.

2.2 Culture Media Preparation

Using the Filter-U.V. system, fill a 20 liter autoclavable Nalgene® container with seawater (clean it before using the same procedure you used for the buckets). Put the lid back on, but make sure to leave it loose so that the container will not explode inside the autoclave. Put autoclave indicator tape on it, and autoclave for 60 minutes. Let the water cool down for at least 24 hours to allow CO2 levels to return to normal. Do not screw the lid back while the water is hot or otherwise the container will implode. Once it has cooled, tighten lid and store in a cool dark place (Fig. 7).

Figure 7. 20 liter Nalgene® container for media sterilization

Immediately before filling the buckets with sterile water from the N algene® container use an Oxford® re-pipettor with a sterile tip (they are inside the oven/incubator kept at 60°C at all times) to add 3.5 ml of each nutrient solution. Use only one tip per solution and container, and after use, place them in a beaker with 10% HCl. Later they will be washed and autoclaved for re-use. Close the container and shake to mix the nutrients. Only prepare enough media to fill all your available buckets. It is not advisable to store seawater with nutrients until the next filling because

To fill the buckets with media take them to Culture Room # 2 and line them up on the table (clean the table with alcohol first). Clean your hands with alcohol and empty the R.O. water present in each bucket into the sink, making sure not to touch the edges of the buckets with your fingers or anything else. When you have six 4 liter containers ready (or three 8 liter containers) remove the lids and place them facing up on a clean surface. Clean your hands with alcohol again as well as the outside surface of the Nalgene® media container. Be very careful not to spill media, pour 3 liters into the bucket (you can see the mark in liters on the side) and move quickly to the next one until you finish all 6 (or all 3 if you are filling 8 liter buckets to the 6 liter mark). Put the lids back and store the buckets next to the hood in Culture Room # 1. With the left over water in the N algene® containers, fill the small clear buckets to the 0.5 liter mark following the same procedure.

2.3 Inoculation

There are three types of inoculations with small-scale cultures: 500 ml, 3 liters, and 6 liters. To inoculate a 500 ml clear bucket, get a back-up stock culture that looks dense, and using the vortex, mix it very well. Clean your hands with alcohol and under the hood pour the test tube culture into the bucket. Close the lid and immediately label the culture remembering to write the date. If the culture is a diatom you have to add two drops of sterile silicate solution to it. As always, use sterile pipettes for this purpose. Place the culture on the culture table until is used to inoculate the next step (Fig. 8).

Figure 8. Small-scale culture table

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 9 4 liter buckets usually have about three liters of media. Before inoculation put a 0.2 urn filter on the air inlet. These filters are unidirectional, so make sure you install them correctly. The filters are always inside the ovenlincubator at 60°C. That should be enough to maintain their sterility (Fig. 9).

Figure 9. 60°C incubator where air filters and pipette tips are kept

Using one of the 500 ml cultures, follow the same inoculation procedure (instead vortexing, just swirl the culture being careful not to spill it). If diatoms are inoculated use 10 drops of silicate solution. 8 liter buckets roughly have 6 liters of media. To inoculate them use also a 500 ml culture following the same procedure. If diatoms are inoculated add I ml of silicate solution. Once cultures are inoculated put them on the table and attach the air line to the filter. Turn the air on until moderate bubbling is going. Too much bubbling will probably kill a fresh inoculum, and weak bubbling will not promote good growth. Until you use them to inoculate carboys, monitor the color to see if they are getting darker.

2.4 Small-scale Culture Table

This table does not have shade screen, however, in some cases, it its good to have some screen in the section where the 500 ml cultures are sitting. Check for bad or blinking bulbs and replace them as necessary. Clean the surface of the table regularly with alcohol.

3.0 Carboy Cultures (16-20 liters)

This is the last stage at which sterile techniques can be used efficiently and cost effectively. Carboys are inexpensive, very resistant to impact and to the autoclaving process. With time they will become opaque and will brake easily, but a $6 dollar carboy can last for over a year even if is subjected to extreme temperatures and pressure once a week.

The culture containers at this stage consist of two elements: The carboy itself, and the air delivery system (I call it a "stem"). Although there are approxim,ately equal number of carboys and stems, the latter are somewhat fragile and are broken when mishandled. Use glass for the stems because it is much more cleaner than plastic, and because it can be autoclaved. The stems consists of a polypropylene, autoclavable 250 ml beaker with a hole on the bottom, which lets through a section of thick medical grade silicon tubing (also autoclavable). At one end, a 35 em piece of glass tubing is attached to the silicon hose. The other end is used for an air filter. Stems should be kept submerged in a chlorine bath at all times. Once they have been used in a culture they are rinsed well and placed in the chlorine bath. This bath is a Tupperware® container with a solution of R.O. water and 200 ml of 10% chlorine. Every two weeks the bath should be emptied and the container cleaned.

There are two types of carboys in our lab. The regular carboys and the "I" carboys (for Inocula) which are used to inoculate the regular carboys. There is no difference between them but they go through different processes so I have marked "I" carboys with red tape around the neck to identify them.

Carboys are cleaned as follows:

• As soon as the culture in the carboy has been used for inoculation of a larger container or other carboys, rinse it with a R.O. water "firehose" blasting the remaining algae attached to the bottom and the walls.

• Add approximately one liter of 10% muriatic acid (l0% HCI) to the carboy and swirl it so the acid touches the whole surface inside. Carboys are left for a minimum of two hours with acid in them to make sure the rernairung algae is killed.

• Rinse carboy with R.O. water. There are two ways to do this: One is using the firehose and the other the "serial washer" which is a 5 outlet firehose that saves a lot of time (Fig. 10) A piece of plywood with 5 holes aligned to the water jets is placed on the sink and the carboys

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 1 1 are turned upside down to receive the blast of water. Carboys should be rinsed for at least a minute to make sure all the acid is removed.

Figure 10. Serial washer

• "I" carboys are washed the same way but being extra careful to get rid of any possible source for contamination. Once they are clean, put one of the chlorinated stems, and a glass plug on the tip of the silicon tubing to prevent spilling during autoclaving. Also, put autoclave indicator tape.

3.2 Culture Media Preparation

Culture media preparation for the carboys follows two methods. Regular carboys are semi-sterilized with chlorine (some spores are able to survive this treatment), while "I" carboys are autoclaved to achieve total sterility.

• Put the regular carboy under the manifold that delivers the seawater (aka "the serial filler", Fig II) and add 10 ml of 10% chlorine. Place one of the chlorinated stems in the carboy and attach the hose from the manifold to the tip of silicon tubing, remembering to open the clamp. Make sure the filters in the fiIter rack are not leaking and are clean. Turn the V. V. columns on and let the system run for several minutes until only seawater is coming out (see "Filtration and V.V. System" appendix). You can also fill each carboy individually.

• Fill the carboys to about 16 liters and detach from the serial filler. Shake the carboy so the chlorinated water touches everything inside. It is advisable to leave the carboy chlorinating for at least 24 hr. but not for more than 6 days (Fig. 12)

Figure 12. Carboy ready to be inoculated

• "I" carboys are filled the same way but skipping the chlorine step. Instead, they are placed on the autoclave "shuttle" and taken to the big sterilizer for one hour. After sterilization, let them cool down for 24 hrs (see "Autoclaving" appendix).

To prepare the media for all carboys use sterile nutrients and an Oxford® pipette with sterile tips (Fig 13.)

Figure 13. Oxford® pipette with ste rile tip (above)

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 1 3 Add 3.0-3.5 ml of each solution (A & B) to the carboy being careful not to touch anything with the tip. Use one pipette tip per solution and place in a beaker with 10% HCI after use. To add nutrients, lift the top of the carboy from the upside down beaker and never touch the edges of the neck. If diatoms are going to be inoculated, add 3 ml of silicate solution to the carboy in the same fashion. Label carboys with masking tape and put a unidirectional 0.2 urn filter on the outside tip of the stem.

3.3 Inoculations

Two kinds of inoculations are performed at this stage. One is the inoculation of "I" carboys using small-scale cultures and the other the inoculation of regular carboys, using "1" carboys.

To inoculate "I" carboys, get a dense 3 or 6 liter culture and carefully pour approximately 1 liter into the carboy. Place carboy on the "Inocula" shelf in Culture Room #1 and connect the air. To inoculate a regular carboy, get one of the dense "I" carboys from the same shelf and split it into 6 regular carboys. To do this lift the stems of the carboys and balance them on one side making sure they do not fall out. In the eventual case this happens, discard that stem and replace it with a clean one. Remove the stem from the "I" carboy and with alcohol and a paper towel, clean the neck. Slowly pour 2.0-3.0 liters of culture into one regular carboy trying not to spill algae, and move to the next without returning the "I" carboy to the vertical position. This is done to prevent the culture from being spilled out of the carboy, touch the neck and then be poured back to a new carboy. Although it seems a bit extreme, this is the only way to avoid contamination. Once the 6 carboys are inoculated put them on the illuminated shelves in rooms #1 or #2 and connect the air to them.

3.4 Culture Room

Carboys are kept on shelves with tluorescent bulbs. Culture room #1 can have up to 77 carboys. 1 have separated the growing section from the inocula section to avoid cross-contamination and confusion. The back wall is also divided in two sections: 1) Tiso (left); 2) Diatoms (right) (Fig. 14)

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 1 4 Figure 14. Carboy arrangement in culture room # 1. The picture on the left shows the Tiso section. The center pictures show the diatom section, and the left picture shows the "Inocula" section

Bulbs that are blinking or are off must be changed as soon as possible. The air system has windows of clear PVC® where you can check for fungi or moisture. If something starts growing inside the airlines they should be bleached. To bleach the airline turn the blower off, disconnect all the carboys from it and close all the valves. Pour a solution of 2% chlorine (dilute 2 parts of 10% chlorine in 8 parts of water) until you see bleach In all the windows. To make sure all air is out of the lines have a bucket at the air line drains and keep pouring bleach solution until all the air is pushed out. Leave the airline bleaching for 4 hours (it is advisable to start this procedure first thing in the morning). After bleaching, the chlorine has to be removed completely from inside the pipes. Even traces of it will kill all the cultures in a matter of minutes. To rinse the bleach you have to connect drain hoses to all the airline drains (4 in total) and connect an R.O. line to one of the inlets (top) (Fig. IS).

Figure 15. The left picture shows the section of pipe used to bleach the airlines. The right picture shows one of the drains at the bottom

Leave the water running for several minutes and then switch inlets and repeat the operation. Check for a change of color in the water, from yellowish to clear, and taste the water to make sure there is no chlorine in it. Turn the blower back on and open all the valves and leave it like that for three hours at least, then connect the carboys back. It would be a good

idea to connect only one carboy and wait for a few minutes to make sure nothing suddenly dies.

Provide CO2 to the cultures to promote growth. The CO2 tank is connected directly to a mixing chamber before the air passes through a 3 urn filter and is distributed to the carboys. I have put a flowmeter to measure the

amount of carbon provided to the cultures. Usually the CO2 flow should be set to 1000-1500 ml min-I, if you need to boost the growth a bit, increase

the flow to 2000 mI min-I but stop there, because too much CO 2 can lower the pH of the media and kill the algae (Fig 16).

Figure 16. Air/Carbon di oxide mixing chamber

Clean the culture room every Friday at the end of the day with floor disinfectant, and wipe the work tables and hood with alcohol.

4.0 Cylinder Cultures in Room #2 (185- 208 liters)

There are two types of cylinder cultures at Harbor Branch. Clean semi sterile cylinders in culture room #2 and non-sterile cultures in the algae greenhouse. The procedures for both are different and will be explained separately.

Culture room # 2 has 16 cylinders and space for 34 carboys. The cylinders have plexiglass lids, silicon tubing, flat bottoms and air filters. In essence, they are just big carboys and should be treated in the same fashion where the culture technique is concerned. The only difference is that they are fixed to a platform and share a common drain.

To prepare a cylinder for culture it first has to be rinsed with R.O. water using a pressure hose (garden hose connected to the R.O. line. If there are residuals from the last culture, remove them using a soft sponge and rinse again. Close the drain and harvest valves and using Alconox soap and a brush clean the inside surface of the tank. Rinse well with R.O., then using the acid sprayer (lO% muriatic) spray the inside of the cylinder making sure every square em is touched by the acid. Use a mask, safety glasses and gloves when you do this (Fig 17.)

Figure 17. Acid sprayer.

Take the lid and silicon tubing (air line) to the sink and nnse them with R.O. water, remove the airstone and put it in a container with acid. Using the sprayer's tip introduce some acid in the tubing and spray both sides of the lid. Rinse them with freshwater and replace the airstone with a dry clean one. Put the lid back on the cylinder and leave the acid for 1 hour inside the cylinder. Open the drain valve and rinse the cylinder with fresh water until all traces of acid are removed. Pay special attention when you do this because there are lights and electrical outlets close to the cylinders and it is not advisable to get them wet. Introduce the tip of the garden hose in the harvest pipe, open the valve and rinse that part of the system. Close all valves and label the cylinder with masking tape and green tape to indicate acid has been used. Leave tank as is, until you are ready to fill it with seawater (Fig. 18).

4.2 Culture Media Preparation

Turn the U. v. lights on and prepare the filtration system (see appendix). Let the filtered seawater run for a while through the pipes until no more R.O. water comes out of it. Wipe the tip (shaped as an "U") of the filtered water delivery hose with alcohol and hang it on the cylinder you are going to fill. Open the valve and let the tube fill (Fig 19).

Figure 19. Cylinder in room # 2 filling

While that is going, add lOO ml of 10 % chlorine solution into the cylinder. Stop the filling when the water gets to very edge of the tube. Leave the tube chlorinating for 24 hr. Make sure the air is off. If the air is left on, the

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 1 8 chlorine will be blown away in a couple of hours and the cylinder will not be sterile the next day. Label the tube with yellow tape to indicate chlorine has been used.

4.3 Inoculation.

To dechlorinate a cylinder, use 15 ml of concentrated sodium thiosulfate (Fri tz® Chlorine Remover, ordered from Aquacenter). Turn the air on for a few minutes to mix the thiosulfate and dechlorinate the whole volume. Check the chlorine levels with GT.G. by taking a small sample from the harvest pipe at the bottom. This is the last place where the thiosulfate will react with the chlorine, so if you get a clean check you are ready to inoculate. If you get color, drain 500-1000 ml from the cylinder and take another sample to check. If dechlorination has not taken place at this point, you may have a bad batch of thiosulfate. Use a new bottle and repeat the procedure. Lower the level to the 200 liter mark by opening the drain valve (Fig 20).

Figure 20. Inocula carboy underneath a ready to inoculate cylinder

Using a sterile 250 ml beaker, pour approximately 35 ml of sterile nutrient solution A and with a 60 cc sterile syringe get the pink liquid and add it to the cylinder. Repeat the operation with solution B (green liquid) and with silicates if you are going to inoculate a diatom. Rinse the beaker and the syringe and put them in the "to autoclave" shelve. Syringes are autoclaved inside an 8 liter clear bucket and beakers are sterilized inside biohazard bags.

Get a dense "I" carboy from underneath the cylinder you want to inoculate.

© Copyright 1996. Harbor Branch Oceanographic Institution Inc . 1 9 Remove the stem. Wipe the outside of the neck with alcohol. Open the cylinder by sliding the lid back a few centimeters. Slowly, trying not to splash, pour the contents of the carboy into the cylinder only touching the edge of it with the clean neck of the carboy. When you are done, slide the lid back and open the drain valve until you return the level of the water to the 200 liter mark. Do this before you turn the air back on, so only unmixed clean water is drained (the culture will be on the upper side of the cylinder). Turn the air on. Label the masking tape with red tape to indicate nutrients have been added, and write the name of the culture (clone) and the date.

4.4 Harvest.

Harvest is achieved by attaching a manifold of three outlets to the harvest drain and having 3 clean carboys at each end receiving the algae. It is best if the whole volume culture is harvested at once. Once harvesting is completed, immediately clean the cylinder before the algae dries on the walls and bottom. Clean any spills you make while harvesting. If you prefer you can harvest one carboy at the time (Fig 21).

Figure 21. The picture on the left shows the common drain shared by the cylinders in room # 2. The right picture shows a cylinder being harvested to 3 carboys

4.5 Culture Room

The culture room # 2 serves a dual purpose. It houses regular cultures (cylinders) and inocula cultures (carboys). Cylinders are labeled from T I to T 16 starting on the left side of the room as you go in. Carboys are not labeled. The ambient temperature in the room should remain at 24-26°C

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 20 while the temperature inside the cultures should read 24-25°C. There is a temperature probe in cylinder T6 which has a digital display right above the tube. Before you leave the room at the end of the day check for bad bulbs and replace them if necessary. Also check that the filtered seawater and R.O. valves are closed. Water pressure can rupture a hose. If unattended, there will be an inundation in the lab. CO 2 and air systems are connected independent from culture room # 1. Check that they are running ok.

5.0 Algae Greenhouse Cultures

In this section even though there are different volume cultures, I will lump everything in one procedure since it is basically the same for all sizes. The main difference with the cultures inside is the level of cleanliness that can be achieved and maintained in the greenhouse. Since it is mainly an "outdoor" operation, cultures tend to get more contamination. Also, the temperature and light parameters are, for the most part, uncontrollable. During the winter months the water gets relatively cold for tropical species like T-iso and since days are shorter, their photosynthetic time is reduced, thus, reducing the growth. At the same time, winter bring less organisms in the water and contamination is easier to control. During the summer months, the temperature raises to intolerant levels in the greenhouses and the water is much more "diverse" in its microfauna and flora, which represents a major problem when trying to minimize contamination. Days are longer during this time and cultures grow very fast but they also crash faster. It is very difficult to control these factors because the system does not allow it. The best thing to do is to be extra careful with the technique and monitor the system often. We are subjected to a number of risks such as the many seasonal ecological successions of our water supply and its regular chemical and salinity changes. There are facts of life that some cultures are going to crash, and that it will be very difficult to pinpoint the cause. Clams WILL be affected by the quality of the algae and since we are producing more than ever, the risk of loosing seed clams due to algae problems is higher. Be prepared for this eventuality.

Cleaning for cylinders, barrels and tanks is basically the same. They are rinsed well with R.O. water, and only when Tiso is going to be inoculated acid, is used as an extra precaution. After the acid, tanks are rinsed again with fresh water. Airstones are changed with each new culture for Tiso, and every 3 culture cycles for the rest of the species (Fig. 22).

Figure barrels and tanks where mi croalgae is cultured in the greenhouse. picture shows portable lights next to a 5' culture barrel

5.2 Culture Media Preparation

For this section I will present the protocol In the form of a table (Table I). Vessels are filled with filtered seawater, chlorinated, dechlorinated, and nutrients are added before inoculation using clean graduated plastic cylinders.

T able I Me d· la prcparation procedure. Vessel Water Filter ChI. Thio. Sol. A Sol. B S i Type Size (u m ) ( m I) ( m I) (m I) (m I) (m I)

Cy Ii nder 3 100 18 30 30 20 Barrel (4' ) 3 (Tiso), 5 (others) 300 100 150 150 80 Barrel (5' ) 3 (Tiso), 5 (others) 400 150 400 400 150 Tank (8 ') 5 500 200 600 600 200

The main concern while preparing the tanks and media for inoculation is to use clean instruments such as pipettes, graduated cylinders, hoses, etc., and to inoculate as soon as the media is prepared. Most culture vessels in the greenhouse do not have covers, or have very precarious lids, which contribute to, or perpetuate contamination problems. It is a priority to

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 22 daily check for fungi and other microorganisms growing in the air and water lines, and if necessary, to bleach those lines on a regular basis.

5.3 Inoculations

To inoculate greenhouse cylinders, carboys from the inside lab are used. They can be regular carboys from Culture Room #1 or carboys harvested from the cylinders in Culture Room #2. Barrels (4') are inoculated using either carboys from inside, or outside cylinders. Barrels (5') are inoculated the same way and tanks (8') are inoculated with barrels. Finally, clams are fed with algae from cylinders, barrels or tanks either by centrifuging (with a continuous centrifuge located outside the algae green house) and collecting a concentrated paste that is later selectively diluted in the clam hatchery, or by pumping the algae to head tanks that will trickle to the clams.

5.4 Greenhouses

The green houses have clear tops to allow light through but there is also artificial illumination that has to be maintained. Check regularly for bad bulbs, and during cloudy days use the portable lights to boost the growth of the cultures. CO2 is very important at this stage and must be checked often. Again, the key to avoid contamination problems is to be on top of the situation, clean everything well, constantly check for suspicious colored spots in all the hoses, and monitor the progress of the cultures frequently. Cleaning the greenhouse on a regular basis is a must when ever time permits (Fig 23).

Figure 23. Greenhouse

Figure 24 summarizes the technique followed at the HBOI Aquaculture Di vision to culture microalgae.

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 23 Microalgae culture process at Harbor Branch Oceanographic Institution - Aquaculture Division .. 4 liter cultures A I ~ ~ I C Filter rack , 25 Jlm Sea water 5 JlIT' ~ ~- ~. rn C . ==Ll ~ 5 urn Stock 250 mililiter 5 urn cult ure cult ure 3 urn

B Sand filter Nutrients (50 11m)

FEEDINGS

H F D =6a,

~ (~~ .'. .,.- ." Algal culture Culture Lab E

Continuous centrifuge G Sterilizat ion (autoclave) Culture Cylinder Culture Tanks Washing (200 liters) (1000-3000 liters)

Figure 24. Microalgae culture process. A) Scaling up cultures; B) Nutrient preparation and addition; C) Sea water filtration and UV exposure; D) Sterilization; E) Culture vessel washing; F) Culture inoculation and m ass algal cultures; G) Culture volum e expantion H) Algae used for feedings, Appendix I

Autoclaving

We have two autoclaves. A small one with capacity for only a few flasks, and an industrial one which can hold up to 10 carboys at once. The instructions for the operation of the small autoclave are written on the front of it. Use this autoclave when you need to sterilize small instruments such as pipettes, test tubes, small tubing, and nutrients. The big autoclave is used almost exclusively for sterilizing carboys.

To operate this autoclave follow these steps:

1- Turn the boiler on

3- Turn power and control switches on . You will get an error message on the display window followed by a print out. Then a beeping sound will follow for a few seconds and then the temperature of the chamber will be displayed. Some times, since the autoclave resides outside and the humidity has ruined some electronic components, the temperature will not be displayed properly.

4- Wait 30-40 minutes for the pressure In the Jacket to raise up to 30 psi. Read this off the gauge.

5- Once the jacket pressure reaches 30 psi, close the door making sure is tight, and turn the water on (blue valve). Leave the two red valves alone. Using the manual controls turn the knob to the "condition" position and wait for 1 minute until the pressure of the jacket decreases to 10 psi.

6- Turn the same knob to the "sterilize" position and time the cycle using the stopwatch. For carboys with filtered seawater or Nalgene®

5- After the time is up, turn the knob to "fast exhaust" and close the blue valve. Steam will rush out of the autoclave exhaust outside the room. Make sure there is nobody standing near by.

6- Wait approximately 30 to 40 minutes for the pressure of the chamber to go all the way down. The autoclave door will not open if the pressure is above O. Nevertheless the security system may fail, so use extra care any time you try to open that door. Remember that the temperature inside is over 124 DC (250 DF).

7- Stay away from the steam that comes out when you open the door and wait a few minutes until the carboys are nor boiling. They have been under pressure and the temperature they supported was above the boiling point of seawater. Some times the water gets into a state called "super heated water" and even though the ambient pressure has return to 1 atmosphere, the temperature of the water remains above the boiling point. The slightest movement will suddenly make the entire carboy to boil in a few seconds. You do not want to be close if that happens. Leave the carboys cooling inside with the door cracked for at least one hour.

8 - Turn controls off and then turn the boiler off.

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 27 9- Using the orange autoclave gloves pull the cart out of the autoclave and drag it back to the lab. Have a sign warning people that the cart is hot.

10 - If there were spills inside the chamber, wait until it IS cold and nnse the inside with a hose.

If the autoclave is not working or if you notice something out of the ordinary, call AMSCO (Tel: 1800-333-8828) Tell the operator that you are referring to the HBOI Aquaculture autoclave. We have a preventive service and maintenance agreement with them.

Filtration and U.V. System

There is a rudimentary filtration system In place at the laboratory. The water comes out of the well and passes through a de-gaser where a myriad of micro organisms thrive. Then the water goes to a reservoir with its own diverse ecosystem. Here also, there is an accumulation of sediment and organic matter which provide surface area for bacteria to grow very nicely. The water is then pumped through a sand filter and then it goes to the laboratory where a series of 5 filters and aU.V. dual column are installed to try to stop those organisms from going into our cultures. After the water goes through these filters, it is distributed to the carboys or cylinders where it is chlorinated or autoclaved. The sand filter has to be back flushed every time you want to use filtered water. To back flush turn the lever to the " back flush" position and wait for 5 minutes until the water coming out of the drain looks clear. Turn the lever to the "rinse" position for one minute and finally turn the lever to the "filter" position. Inside the lab, prepare the filter rack by putting a set of clean filters.

When you are ready to use the water turn the V.V. system on and open the well water valve and the by-pass valve but keep the filter rack valve closed. This way the pipes can be purged without clogging the filters. Once the water comes out clean, close the by-pass valve and open the filter rack

© Copyright 1996. Harbor Branch Oceanographic Institution Inc. 29 valve. Usually these filters are sitting In chlorinated R.O. Water so you have to let the well water run for a few minutes. You can use this water in three ways: 1) Use the extension hose to fill carboys that are not sitting under the serial filler, to fill Nalgene® containers for the preparation of media, or carboys in the autoclave shuttle. 2) Use the serial filler to fill 45 carboys at once. 3)- Use the extension hose in culture room #2 to fill the cylinders.

After any of these uses, you have to back flush the pipes with R.O. water so nothing grows in them until the next filling. For that purpose, open all the valves in the system and connect the R.O. hose to the extension hose you used. Open the by-pass valve and turn the R.O. water on. Wait a few minutes until only fresh water is coming out. You can then pour 60 ml of chlorine inside the R.O. Line and turn it on again. This will take high concentrations of chlorine to the filters. As soon as you smell the chlorine coming out of the by-pass pipe turn to R.O. water off and close all the valves.

Clean the filters with Micro® every other filling or alternate the filters each filling with a new or clean. set. One set of filters should no be used more than 10 times

Appendix III

Thiosulfate and Silicates

We use two kinds of thiosulfate solutions. One we do not have to prepare because we buy it from Aquacenter. It is called "Chlorine Remover" and it IS very concentrated. We use chlorine remover for the dechlorination of carboys and cylinders. The other type of thiosulfate we prepare from an industrial powder and it is not as concentrated. We use it for barrels and tanks. To prepare this solution dissolve 250 gm of the powder into I liter of water on a magnetic stir plate.

Silicates are used in the diatom cultures because diatoms build a hard silicate shell around their cell wall. The silicate solution is prepared dissolving 200 gm of sodium metasilicate in one liter of water. You will need a magnetic stirrer and a hot plate to help the dilution processes.